Download Bacterial Death Results from Mutations Made in the Translocation Peptide... tRNA Synthetase

Bacterial Death Results from Mutations Made in the Translocation Peptide of LeucyltRNA Synthetase
Daniel M. McDonough, Layla N. Baykal, Mallory J. Banton, and Rachel A. HellmannWhitaker
Department of Chemistry and Physics
Coastal Carolina University
ABSTRACT
The family of aminoacyl-tRNA synthetases (aaRSs) ensures the fidelity of translation
through providing a pool of correctly aminoacylated tRNA products that become
incorporated by the ribosome. Leucyl-tRNA synthetase (LeuRS) has two functionally
separate domains, one is the aminoacylation domain and the other is the CP1 editing domain.
LeuRS can aminoacylate noncognate amino acids, therefore it relies on the CP1 editing
domain to hydrolyze misaminoacylated tRNA products before they are released from the
enzyme. The LeuRS enzyme must undergo a structural transition state in its reaction cycle in
order to translocate the 3’ acceptor stem of tRNA 30 Å from the aminoacylation active site to
the CP1 domain hydrolytic active site. The translocation event is difficult to study, but we
believe that we have generated mutations within LeuRS that alter the translocation event of
tRNA. The mutations that we have generated lead to bacterial death in Escherichia coli (E.
coli). Circular dichorism experiments indicate that our mutations do not significantly alter the
secondary structure of LeuRS. In vitro biochemical studies demonstrate that these mutations
reduce the rates of aminoacylation and hydrolysis, while also displaying misaminoacylation
activity. We attribute these biochemical findings to the resulting bacterial death that is caused
by these mutations.
Introduction
Polymerization of amino acids during canonical protein translation is an accurate process
with a misincorporation rate of 1 per 2000 amino acids (Drummond & Wilke, 2008). During
translation, cells employ many quality control mechanisms that maintain the precise
production of proteins. The first quality control step is maintained by aminoacyl-tRNA
synthetases (aaRSs) through the synthesis of aminoacylated tRNAs. All aaRSs rapidly bind
their cognate amino acid and tRNA to turn-over an accurately aminoacylated tRNA product
(Hendrickson & Schimmel, 2003). To ensure the fidelity of aminoacylation many aaRSs rely
on a two-active site system termed “the double sieve model” for editing mistakes (Fersht,
1998; Fersht & Dingwall, 1979). Aminoacylation takes place in the synthetic active site
termed “the coarse sieve,” which can exclude structurally dis-similar substrates. The second
sieve is termed “the fine sieve,” and acts through a hydrolysis reaction to cleave incorrectly
aminoacylated homologous products. When the editing process has been rendered defunct
through mutational analysis, the results have been found to be deleterious for both
prokaryotic and eukaryotic cells (Karkhanis, Boniecki, Poruri, & Martinis, 2006; Lee et al.,
2006; Nangle, Zhang, Xie, Yang, & Schimmel, 2007)
The family of aaRSs are divided into two classes that are believed to have co-evolutionary
pathways that have functionally merged from two different primitive biological
processes(Woese, Olsen, Ibba, & Soll, 2000). Class I and class II aaRSs are distinct in both
their structural features and their respective mechanisms of aminoacylation(Ibba & Soll,
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2000; Woese et al., 2000). These structural features are known as protein folding motifs,
which commonly occur in nature. Once such structural motif is the Rossmann fold, which is
known to bind to nucleotides and nucleotide derivatives such as nicotinamide adenine
dinucleotide (NAD)(Rao & Rossmann, 1973). Class I aaRSs employ the Rossmann fold to
house the aminoacylation active site, which structurally consists of parallel - strands with
the signature amino acid sequences KMSKS and HIGH (Rould, Perona, Soll, & Steitz, 1989;
Schmitt, Meinnel, Blanquet, & Mechulam, 1994; Webster, Tsai, Kula, Mackie, & Schimmel,
1984). The active site within the Rossmann fold binds to ATP, the cognate amino acid and
tRNA to carry out the two-step aminoacylation reaction (as can be seen below in the reaction
schematic) (Arnez & Moras, 1997). For some class I aaRSs there is a second active site,
which is housed within a separate domain. To generate this second domain, the Rossmann
fold is split by a connective polypeptide domain (CP1) insertion sequence (Cusack,
Yaremchuk, & Tukalo, 2000). The CP1 domain is only present in 4 of the 11 class I aaRSs.
The CP1 domain houses the editing active site, which structurally excludes correctly
aminoacylated products while hydrolyzing misaminoacylated amino acids from the tRNA
(Mursinna, Lincecum, & Martinis, 2001; Swairjo & Schimmel, 2005). These two actives sites
work in conjunction to ensure that a pool of viable aminoacylated tRNAs are ready for
ribosomal incorporation during protein translation, thereby reducing the misincorporation rate
(Hendrickson & Schimmel, 2003).
Optimal coordination of the two active sites requires communication. The charged tRNA
product initially within the aminoacylation active site, chemically communicates with the
CP1 domain as it translocates into the editing hydrolytic active site (Hellmann & Martinis,
2009). In addition, there is structural communication that requires the CP1 domain to toggle
in relation to the aminoacylation active site via flexible linker sequences that act as a
molecular hinge (Mascarenhas & Martinis, 2008). These processes have been studied
extensively for the class I aaRSs that have the CP1 domain insert such as: leucyl-tRNA
synthetase (LeuRS) and the homologous isoleucyl-tRNA synthetase (IleRS) and valyl-tRNA
synthetases (ValRS) (Starzyk, Burbaum, & Schimmel, 1989). Interestingly, though the two
domains have many modes of communication, the CP1 domain is known to fold
independently of the aminoacylation domain (Cusack et al., 2000; Fukai et al., 2000; Silvian,
Wang, & Steitz, 1999). In addition, when the aminoacylation and CP1 domains have been
expressed as separate independent enzymes, they can still carry out their respective reactions
(Betha, Williams, & Martinis, 2007; Boniecki & Martinis, 2012; Lin, Hale, & Schimmel,
1996; Zhao et al., 2005).
X-ray crystallography data has recently shown that E. coli LeuRS’s structure during the
reaction cycle is highly dynamic (Palencia et al., 2012). Many of the domains undergo drastic
structural and positional changes as the enzyme initiates aminoacylation followed by the
hydrolytic reaction (Palencia et al., 2012). These structural and positional changes help
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support the movement of tRNA between the aminoacylation and editing active sites, which
are separated by about 30 Å (Palencia et al., 2012). It has been previously reported that the 3’
acceptor stem of the tRNA must be translocated between the two active sites during LeuRS’s
reaction cycle (Lincecum et al., 2003; Mascarenhas & Martinis, 2008). Furthermore, the
tRNA’s translocation is believed to be specifically supported by interactions with the CP1
domain (Hellmann & Martinis, 2009; Mascarenhas & Martinis, 2008).
We hypothesize that the tRNA 3’ acceptor stem has to first be translocated into the
aminoacylation active site before the enzyme can begin its reaction cycle and that the CP1
domain structurally supports this initial translocation event, as has been previously suggested
(Rock et al., 2007). Two computationally predicted hinge sites have been identified in the
CP1 domain, which suggests that this region of the enzyme is highly dynamic and therefore
capable of supporting this type of movement (Hellmann & Martinis, 2009; Mascarenhas &
Martinis, 2008). The two molecular hinge sites have been previously described as the strand linker sequences and the translocation peptide (Hellmann & Martinis, 2009;
Mascarenhas & Martinis, 2008). Additional mutations (D391A and F382A/N383A (FN1))
have been identified in the translocation peptide that has rendered the LeuRS enzyme nonfunctional in E. coli cells. Herein, we report our biochemical findings to support the claim
that these additional sites within the translocation peptide may be integral in the preaminoacylation and post-aminoacylation movements of tRNA into and out of aminoacylation
active site. These mutations have been found to reduce the rates of aminoacylation,
hydrolysis, and to have misaminoacylation activity. Due to the reduced chemical reaction
rates, we believe that the mutations may play a role in stabilizing the tRNA as it translocates
between the two active sites.
Materials and Methods
Materials
T7 RNA polymerase (Invitrogen, Carlsbad, CA) was used to in vitro transcribe tRNALeu via
run-off transcription (Sampson & Uhlenbeck, 1988). The in vitro transcription protocol is as
follows: transcripts encoding the tRNALeu gene were obtained first by purifying 450 g/mL
of ptDNAleu14 (Normanly, Ogden, Horvath, & Abelson, 1986) using an alkaline lysis
protocol. The ptDNAleu14 was restricted digested with BstNI and incubated overnight at 60
o
C. Digested plasmid was confirmed by agarose gel electrophoresis. The product resulting
from the BstNI digest was used as a template for in vitro transcription reactions containing 40
mM Tris-HCl, pH 8.0, 30 mM MgCl2, 5 mM dithiothreitol (DTT), 0.01 % Triton X-100, 40 U
RNase inhibitor (Eppendorf, Westbury, NY) 8 g/ml pyrophosphatase (Sigma, St. Louis,
MO), 5 mM spermidine, 50 g/ml bovine serum albumin (BSA), 7.5 mM each of ATP, CTP,
GTP, and UTP, 80 mg/ml polyethylene glycol (PEG) 8000, 60 g/ml DNA plasmid template
and 17 M T7 RNA polymerase. The tRNA product was purified by urea-containing
polyacrylamide gel electrophoresis (Sampson & Uhlenbeck, 1988) and quantitated based on
absorbance at 260 nm using an extinction coefficient of 840,700 L/mol●cm (Puglisi &
Tinoco, 1989). Purified tRNALeu was denatured at 80o C for 1 min, followed by an addition of
1 mM MgCl2 and quick-cooling on ice for re-folding.
Methods
Generating Mutations: The wild type E. coli LeuRS gene is encoded within the plasmid
pRWecLeuRS, which was generated by extracting E. coli genomic DNA and utilizing PCR to
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amplify and subsequently clone leuS (GeneBank Accession #EG10532) into pET15b (Fisher
Scientific, Pittsburg, PA) as previously demonstrated by Härtlen, M. and Madern, D.,
1987(Hartlein & Madern, 1987). To generate pRWecLeuRS we used synthesized
oligonucleotides (Integrated DNA Technology (IDT), Coralville, IA). The forward primer 5’CGCCATATGATGCAAGAGCAATACCGCCCG-3’
and
reverse
primer
5’CGCGGATCCCGGTTGCTGGTCTAACTCCTC-3’, which contain the NdeI and BamHI
restriction sites, respectively. The resulting pRWecLeuRS was used to insert mutations via
PCR generating the plasmids: pRWFN1 (F382A/N383A), and pRHD391A (D391A). These
mutations were constructed using synthesized complementary oligonucleotides, the FN1
mutation
was
generated
with
the
forward
primer
5’GACTGAAAAAGGCGTGCTGGCTGCATCTGGCGAGTTCAACG-3’ and the D391A
mutation
was
generated
with
the
forward
primer
5’CGAGTTCAACGGTCTTGCCCATGAAGCGGCCTTCAACG-3’.
In
addition,
we
generated the T252Y mutant to purify the mischarged product [3H] Ile-tRNALeu (as described
below) with the following forward primer: 5’-CCCGCCGGACTACTTTATGGGTTGTACC3’.(Mursinna & Martinis, 2002) DNA sequencing for all plasmids was carried out by
Eurofins MWG Operon, Huntsville, AL.
Protein Expression: To express the LeuRS mutants FN1 and D391A, the pRHFN1 and
pRHD391A plasmids were used to transform E. coli BL21 (Stratagene, La Jolla, CA). A
single colony was used to inoculate 5 ml of Luria-Bertani medium containing 100 g/ml
ampicillin (LB-Amp) and grown overnight at 37 oC. The overnight culture was then
transferred to 1 L of LB-Amp and grown at 37 oC to an OD600 of 0.6-0.8. Expression of
LeuRS was induced with 1 mM IPTG for 3 hrs at 37 oC. Harvested cells were lysed by
sonication and each LeuRS, which contained an N-terminal six-histidine tag, was purified by
affinity purification using HIS-Select HF nickel affinity resin as described before
(Mascarenhas & Martinis, 2008). The purified proteins were concentrated via Pierce
concentrators 50K MWCO (Thermo Scientific, Rockford, IL) and quantitated using the BioRad protein assay according to the commercial protocol.
Growth Curve Analysis: Growth curves for selected E. coli KL231 transformants that
contained plasmids pRWFN1 and pRWD391A were grown in 3 ml of minimal standard (MS)
medium (Low, Gates, Goldstein, & Soll, 1971). The MS-media was supplemented with 100
µg/ml of each of the following amino acids: leucine, isoleucine, valine, arginine, histidine,
lysine, methionine, phenylalanine, threonine, tryptophan, and tyrosine. It also included 40
µg/ml uracil, 40 µg/ml adenine, 200 µg/ml thymine, 100 µg/ml thiamine, and 2% glucose.
Antibiotics including kanamycin (30 µg/ml), streptomycin (50 µg/ml), and ampicillin (100
µg/ml) were also added to generate MS-Amp/Kan/Str. Overnight cultures were transferred to
50 ml of MS-Amp/Kan/Str medium for growth curve analysis, which was conducted at 42 °C.
Aliquots of 1 mL were taken every 2 hr until 20 hr of incubation were completed. At 30 hr
and 48 hr another set of aliquots were recovered. Aliquots of the cell cultures were analyzed
by taking the OD600 using an ultraviolet-visible spectrophotometer (Thermo Scientific,
Waltham, MA).
Growth curves were also performed in E. coli BL21 cell (Stratagene, La Jolla, CA)
transformants of pRWFN1 and pRWD391A. A single colony was used to inoculate 5 ml of
LB-Amp and grown overnight at 37 oC. Overnight cultures were transferred to 50 ml of LBAmp for growth curve analysis, which was conducted at 37 °C. Aliquots of 1 mL were taken
every 2 hr until 20 hr of incubation were completed. At 30 hr and 48 hr another set of aliquots
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were recovered. Aliquots of the cell cultures were analyzed by taking the OD600 using an
ultraviolet-visible spectrophotometer (Thermo Scientific, Waltham, MA).
Aminoacylation Assays: Leucylation reactions contained 60 mM Tris-HCl, pH 7.5, 10 mM
MgCl2, 1 mM DTT, 4 M folded tRNALeu, 21 M [3H]-L-leucine (150 Ci/ml) and 50 nM
enzyme. Alternatively, isoleucylation reactions contained the same components, except 21
M [3H]-L-isoleucine (radioactivity) was added instead of the tritiated leucine. Initial
velocities for the leucylation reactions were obtained using 10 nM of enzyme. All reactions
were initiated by the addition of 4 mM ATP and then quenched at specific time points by
transfer to filter pads (Whatman, Clifton, NJ) that had been pre-soaked with 200 L of 5%
trichloroacetic acid (TCA). The pads were then washed three times with cold 5% TCA,
followed by one wash with cold 70% ethanol at 10 min time durations for each washing. The
pads were dried by soaking in ethyl ether and then under an ultra-red lamp. Each pad was
transferred into a 20 ml scintillation vial with 3 mL of scintillation fluid (Fisher Scintisafe
EconoF) and quantitated using a HIDEX Triathler Liquid Scintillation Counter.
Charged tRNA Preparation and Enyzmatic Deacylation: The [3H]-Leu-tRNAleu was
generated in a one hour incubation of the leucylation reaction described above. The [3H]-IletRNAleu was prepared using an editing defective T252Y LeuRS mutant in an aminoacylation
reaction that was incubated at 30 oC for 3 hr. Reactions were stopped using 0.18% acetic
acid, and extracted using two equal volumes of phenol/chloroform/isoamyl alcohol, pH 4.3
(125:24:1) (Schreier & Schimmel, 1972). A one-half volume of 4.6 M NH4OAc, pH 5.0, was
added followed by an ethanol precipitation. The dried aminoacylated tRNA pellets were
resuspended in 10 mM KH2PO4, pH 5.0. Hydrolytic editing assays were carried out at room
temperature in 60 mM Tris, pH 7.5, 10 mM MgCl2, and 0.8 M [3H]-Leu-tRNALeu or 0.8 M
[3H]-Ile-tRNAleu and was initiated with 100 nM enzyme (Mursinna et al., 2001). At specific
time points, 5 l aliquots were spotted onto filter pads, washed, and quantitated.
Structural Analysis of Wild type, FN1 and D391A mutants: Circular Dichorism was used to
characterize the relative -helix and -sheet content of the wild type and mutant LeuRS
enzymes. Each enzyme was diluted to 0.8 mg/mL in potassium phosphate buffer (10 mM
KH2PO4 and 10 mM K2HPO4 pH 7.4). The enzymes were then inserted into a JASCO J-810
circular dichroism spectropolarimeter set at 25 oC using 50 nm/min scan speed, 0.5 sec time
constant and a 1 nm bandwidth (Kelly, Jess, & Price, 2005).
Results
Examination of the Structural Impact of the FN1 and D391A mutations
The LeuRS mutations FN1 and D391A reside within the translocation peptide. The
translocation peptide is approximately 12 Å away from bound tRNA (Hellmann & Martinis,
2009). Based on x-ray crystallography images of LeuRS in the aminoacylation and editing
conformations, the translocation peptide is projected to move ± 12-15o with the rotational
movement of the CP1 domain (Figure 1A) (Palencia et al., 2012).
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Figure 1A
Figure 1A. Secondary and Tertiary structure of the E. coli LeuRS and FN1 and D391A Mutants. E. coli LeuRS
aminoacylation and editing X-ray crystal structures (Palencia et al., 2012). The CP1 domain (yellow) is inserted into the
catalytic aminoacylation core (green) via two β-strand linkers. The mutants D391A and FN1 are depicted as ball and stick
structures colored blue.
Previously published results have shown that the residues H392 and E393 specifically
interact with the tRNA molecule to promote its translocation between the two active sites of
LeuRS. This claim was supported by biochemical and computational evidence (Hellmann &
Martinis, 2009). The residues D391, H392 and E393 are all part of a computationally
predicted hinge region that has been projected to be a very dynamic region of the CP1
domain. As can be seen in figure 1A, the molecular hinge divides the translocation peptide
into two halves, with both ends forming -helical structures. D391 sits in the center of
translocation peptide. Being that D391 is at the center of the molecular hinge site, it may
contribute to crucial conformational changes the translocation peptide may experience during
the transient translocation step. In addition, because of the position of D391, this residue
could form electrostatic interactions with the bound tRNA. The F382 and N383 residues are
outside of the molecular hinge region, and are at the distal end of the translocation peptide in
relation to tRNA (Figure 1A). Therefore, it would be unlikely that these residues have direct
interactions with tRNA. However, based on the structures of LeuRS and the positions of
F382 and N383, these residues most likely play vital secondary structural roles in helping to
provide the correct orientation of the translocation peptide so that it may interact with the
tRNA. F382 and N383 are proximal to other aromatic residues in the CP1 domain, which
could allow for base-stacking interactions thus stabilizing the translocation peptide.
As will be addressed, these two mutations have been characterized as non-functional in E.
coli cells. To rule out the possibility that the bacterial death caused by these mutations result
Bridges 9 (Spring 2015)
6
from regional or global denaturation of the secondary structural features of LeuRS, we
conducted circular dichorism analysis of the mutant enzymes in relation to the wild type
enzyme (Figure 1B). As indicated by figure 1B, the circular dichorism spectrum indicates
that the mutants have similar secondary structural features to the wild type enzyme.
B.
/d m o l)
15000
W T
FN1
10000
D 391A
[  ] (d e g . c m
2
5000
0
200 210 220 230 240 250 260 270 280 290 300
-5 0 0 0
W a v e le n g t h ( n m )
-1 0 0 0 0
Figure 1B
Figure 1B. Secondary and Tertiary structure of the E. coli LeuRS and FN1 and D391A Mutants. Circular Dichorism
spectrum of WT (wild type), ( ); FN1, ( ); and D391A, ( ). Sample concentration was 0.8 mg/mL in potassium phosphate
buffer (10 mM KH2PO4 and 10 mM K2HPO4 pH 7.4). Each enzyme was scanned in triplicate using a JASCO J-810 circular
dichroism spectropolarimeter set at 25 oC using 50 nm/min scan speed, 0.5 sec time constant and a 1 nm bandwidth (Kelly et
al., 2005).
Growth curve analysis indicates FN1 and D391A are non-functional in E. coli cells
The pRWFN1 and pRWD391 plasmids were used to transform E. coli BL21 cells and were
then grown in liquid LB-Amp media over a period of 48 hrs. The growth of the cells was
monitored spectroscopically to determine their intracellular behavior. The FN1 and D391A
mutants grew in a similar manner to the wild type enzyme (Figure 2A).
O .D . 6 0 0
1 .8
A.
W T
1 .5
FN1
1 .2
D 391A
0 .9
0 .6
0 .3
0 .0
0
10
20
30
40
50
60
T im e ( h r s )
Figure 2A
Figure 2A. Growth curve of wild type LeuRS, and the mutants FN1, and D391A, using E. coli BL21 strain and the
temperature sensitive E. coli KL231 strain. Growth Curve of WT, FN1 and D391A in E. coli BL21. Cell growth was carried
out at 37 oC. Symbols represent the following: WT (wild type), (■); FN1, (▲); and D391A, (◊). Error bars are present for all
points and represent growth curves that were repeated in triplicate.
Normal growth was observed because the genomic copy of wild type LeuRS was still active
in the E. coli BL21 cells, thus rescuing growth. Interestingly, we observed a similar result
when the pRWFN1 and pRWD391 plasmids were used to transform the temperature sensitive
E. coli KL231 cells, which were grown at 30 oC. Again, this result is due to expression of
genomic LeuRS within E. coli KL231 cells at 30 oC (Low et al., 1971). The non-functional
behavior of these mutants became evident when the genomic copy of LeuRS was rendered
functionally inactive within E. coli KL231 cells grown at 42 oC (Figure 2B) (Low et al.,
1971).
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1 .8
B.
W T
O .D . 6 0 0
1 .5
1 .2
0 .9
0 .6
D 391A
0 .3
FN1
0 .0
0
10
20
30
40
50
60
T im e ( h r s )
Figure 2B
Figure 2B. Growth curve of wild type LeuRS, and the mutants FN1, and D391A, using E. coli BL21 strain and the
temperature sensitive E. coli KL231 strain. Growth Curve of WT, FN1 and D391A in E. coli KL231. Cell growth was
carried out at 42 oC. Symbols represent the following: WT (wild type), (■); FN1, (▲); and D391A, (◊). Error bars are
present for all points and represent growth curves that were repeated in triplicate.
At this elevated temperature, both mutants were unable to complement E. coli KL231 as
shown in Figure 2B and therefore failed to significantly support cellular growth. Therefore,
from the growth curve analysis, we have shown that FN1 and D391A reduce the growth of E.
coli cells by approximately 80% and that this finding indicates that these mutations are nonfunctional. Thus, these mutations must significantly impact the biochemical behavior of
bacterial LeuRS.
Biochemical evidence supports the role of FN1 and D391A in tRNA movement
In order to determine how these enzymes impact the biochemical behavior of LeuRS, the
FN1 and D391A mutants were purified through ion-exchange chromatography as previously
described in the Methods section. The purified enzymes were initially tested for their ability
to leucylate tRNA (Figure 3A).
L e u -tR N A
Leu
( p m o l)
20
A.
18
16
W T
14
FN1
12
10
8
D 391A
6
4
2
0
0
5
10
15
T im e ( m in )
Figure 3A
Figure 3A. Biochemical Analysis of WT, FN1 and D391A LeuRS Enzymes. Leucylation activity of wild type and mutant
LeuRSs. The reaction was carried out with 50 nM LeuRS, 4 μM tRNALeu transcript, and 22 M [3H]-leucine (167 Ci/ml).
Symbols represent: WT (wild type), (■); FN1, (▲); D391A, (◊); and No Enzyme (x). Error bars are present for all points and
represent reactions that were repeated in triplicate.
As indicated in figure 3A, these mutants have reduced leucylation activity when compared to
the wild type enzyme. This finding correlates with the results of bacterial death by suggesting
that the mutant enzymes cannot turn-over leucylated tRNA fast enough for ribosomal
incorporation. This would then lead to leucine-deficient proteins and thereby increase the
amount of aberrant proteins being produced in the cell, thus cellular death.
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Additionally, it can be seen in figure 3A that both FN1 and D391A fail to reach a maximal
reaction velocity, which is represented by the plateau of leucylation activity (as seen in the
wild type results). This plateau value represents an equilibrium between the acylation event
and the deacylation events (Bonnet & Ebel, 1972). The deacylation events can occur through
spontaneous hydrolysis of the ester linkage between the amino acid and tRNA or through
enzyme-mediated deacylation, which also results in the hydrolysis of the acylated product
(Bonnet & Ebel, 1972). For the wild-type enzyme these processes are in near exact
equilibrium, but for FN1 and D391A equilibrium is never reached (figure 3A). The results
indicate that the mutant enzymes may reduce the rates of translocation and therefore fail to
produce enough acylated product to be in equilibrium with the deacylated products.
Based on our data, FN1 and D391A seem to influence the translocation of the 3’ acceptor
stem of tRNA into the leucylation active site and then into the CP1 editing active site. We
preformed deacylation assays with the misacylated substrate Ile-tRNALeu as previously
described. In relation to wild type LeuRS, FN1 and D391A had very slow deacylation
activity and failed to reach an equilibrium state (Figure 3B).
B.
100
% Ile - t R N A L
eu
No
E n zym e
80
D 391A
FN1
60
40
W T
20
0
0
2
4
6
8
10
T im e ( m in )
Figure 3B
Figure 3B. Biochemical Analysis of WT, FN1 and D391A LeuRS Enzymes. Deacylation of mischarged E. coli Ile-tRNALeu
by wild type and mutant LeuRSs. Reaction conditions include 1.5 µM [ 3H]-Ile-tRNALeu and 5 nM enzyme. Symbols
represent: WT (wild type), (■); FN1, (▲); D391A, (◊); and No Enzyme (x). Error bars are present for all points and
represent reactions that were repeated in triplicate.
FN1 and D391A deacylated approximately 50% of the available Ile-tRNALeu substrate. In
general, deacylation activity is represented by three events that cannot be separated in this
assay. First, there is spontaneous deacylation occurring in solution (Bonnet & Ebel, 1972).
Second, there is enzymatic deacylation which involves the translocation of tRNA into the
editing active site (Hellmann & Martinis, 2009). Third, is the deacylation of tRNA when it
binds directly in the editing active site without undergoing a translocation event (Betha et al.,
2007). Therefore, from our data it appears that FN1 and D391A both have significantly
reduced translocation activity of the 3’ acceptor stem of the tRNA into the editing active site.
Lastly, we conducted a misaminoacylation biochemical assay to assess whether FN1 and
D391A would allow for the misaminoacylation of tRNA in the presence of noncognate amino
acid. We have likened the aminoacylation active site to a “coarse sieve” whereby all
structurally homologous amino acids would be charged to the 3’ acceptor stem of tRNA. The
FN1 and D391A mutants are believed to inhibit efficient tRNA translocation between the
active sites. Figure 3C indicates that FN1 and D391A have misaminoacylation activity in the
presence of noncognate amino acid. From the graph, FN1 and D391A have 2.5-5 times
higher misaminoacylation activity than the wild type enzyme. These mischarging activities
are directly correlated to a defunct translocation pathway resulting from these two mutations.
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Figure 3C
Figure 3C. Biochemical Analysis of WT, FN1 and D391A LeuRS Enzymes. Misaminoacylation activity of wild type and
mutant enzymes. Isoleucine mischarging reactions were carried out with 1 M LeuRS enzyme, 4 μM tRNALeu transcript, and
22 M [3H]-isoleucine (167 Ci/ml). Symbols represent: WT (wild type), (■); FN1, (▲); D391A, (◊); and No Enzyme (x).
Error bars are present for all points and represent reactions that were repeated in triplicate.
Discussion and Conclusions
From the structural and biochemical data we have gathered, FN1 and D391A do not appear to
significantly impact the structural or functional characteristics of either active site of LeuRS.
Structural data obtained through circular dichorism suggests that these mutants do not
significantly alter the regional or global structure of LeuRS when compared to the spectra of
the wild type enzyme. Furthermore, based on recently published x-ray crystallography data
(Palencia et al., 2012), we believe these sites of mutations are too far removed from either
active to significantly impact their functionality. Since we can rule out the possibility that
these mutants structurally alter LeuRS, we believe that these mutants influence the
biochemistry of LeuRS by altering the chemical communication between the active sites.
We hypothesize that FN1 and D391A affect the chemical communication between the two
active sites by inhibiting efficient movement of the tRNA. Based on our findings, we have
demonstrated that both of these mutations reduce the rate of leucylation as well skew the
equilibrium of product formation. It has been previously reported that the leucylation reaction
is a two-step reaction which can be broken down into a synthesis step followed by a productrelease step (Francklyn, First, Perona, & Hou, 2008; Hellmann & Martinis, 2009). Mutations
made within the translocation peptide were reported to alter the rate of the product-release
step (Hellmann & Martinis, 2009). By extension, we hypothesize that D391A (which is
within the molecular hinge region) and FN1 alter the rate of leucylation as was demonstrated
in figure 3A, by reducing the efficiency of the tRNA translocation.
Based on the biochemical evidence presented in figure 3A, we hypothesize that the reduction
in leucylation activity is two-fold. First, the initial rates of leucylation for the mutant enzymes
are reduced by 50% or more. It has been previously proposed that tRNA translocation
consists of the initial movement of the tRNA 3’ acceptor stem into the leucylation active site
and then a second translocation event which places the 3’ acceptor stem in the hydrolytic
active site (Hellmann & Martinis, 2009; Rock et al., 2007). If FN1 and D391A play a
structurally supportive role in the translocation peptide’s interaction with tRNA, then the
initial rates of leucylation would naturally be reduced as the CP1 domain is less effective at
initiating the movement of tRNA into the leucylation active site. Thus, this biochemical data
suggests that FN1 and D391A both may cause the early release of tRNA before it is
translocated to the CP1 domain for editing. The early release of the tRNA may happen before
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the tRNA is charged with amino acid during the initial translocation into the aminoacylation
active site.
As a result of this hypothesis, we propose that these mutants may initiate a leaky futile
reaction cycle. The futile reaction cycle may consist of the following steps: 1. LeuRS binds
tRNA, 2. enzyme fails to rapidly aminoacylate the tRNA, 3. LeuRS pre-maturely releases the
tRNA, 4. the tRNA re-binds to the enzyme. This futile reaction mechanism would
significantly reduce the population of leucylated tRNA in the reaction tube, thus increasing
the concentration of deacylated tRNA. Therefore, the equilibrium of the leucylation reaction
is skewed toward the deacylated product, which can be seen in figure 3A as both mutants fail
to reach the maximal reaction velocity of wild type LeuRS.
We have hypothesized that FN1 and D391A affect the pre-aminoacylation translocation step,
but we also believe these mutants may affect the post-aminoacylation translocation step. Our
biochemical data supports this finding. In figure 3B, the rates of hydrolysis for both mutants
are reduced, which we hypothesize is the result of a defunct translocation pathway into the
editing active site. The hydrolysis activity that was observed is mainly attributed to the direct
binding of the mischarged tRNA into the editing active site without undergoing translocation.
We have hypothesized that the hydrolysis reaction encompasses non-enzymatic hydrolysis,
hydrolysis via direct binding to the editing active site and hydrolysis through a translocation
pathway that originates at the leucylation active site and ends at the editing active site. If all
modes of editing were occurring we would see the rapid hydrolysis of misacylated tRNA
until equilibrium was reached, as is the case with wild type LeuRS. However, for FN1 and
D391A we see a reduced rate of hydrolysis as well as a failure to reach equilibrium. We
believe that this result stems from a lack of translocation of tRNA between the two active
sites, therefore the only operating modes of hydrolysis are non-enzymatic and the direct
binding of tRNA to the CP1 domain.
Additionally, this finding is supported by the misaminoacylation activity of both FN1 and
D391A. We have shown (Figure 3C) that these mutants produce approximately 2-5
picomoles of Ile-tRNALeu in a 90 minute reaction. This product is less than half of the LeutRNALeu product that is generated by these mutants in a 15 minute reaction. Though these
mutants do not have robust mischarging rates, we believe that this finding is significant. We
attribute the formation of aberrant product to the lack of movement of tRNA into the CP1
domain from the aminoacylation active site. Just like with previous biochemical findings, we
feel that the initial rates of translocation into the active site are reduced, thus dropping the
overall production of product. Once mischarged product is generated, it is released from the
enzyme without undergoing translocation, which allows mischarged product to be detected.
Non-enzymatic deacylation and deacylation via direct binding to the CP1 domain also occur,
which further reduces the concentration of mischarged tRNA product.
From these in vitro studies, we have advocated that our mutants have defunct translocation
pathways for both the pre-aminoacylation and post-aminoacylation translocation events. We
have also supported our findings with in vivo experiments which demonstrate that FN1 and
D391A are non-functional in E. coli cells. We hypothesize that these mutants reduce the rates
of the canonical biochemical reactions of LeuRS and thus render the mutant enzymes unable
to produce sufficient product to meet the synthesis demands of the bacterial cell. When
expressed in bacterial cells, these mutant enzymes may result in a reduced occurrence of
ribosomal leucine incorporation during translocation.
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In conclusion, we hypothesize that these mutations have not significantly altered the structure
of LeuRS. We further believe that these mutations do not directly alter the functionality of the
active sites. Based on our data, we believe that these residues in the wild type enzyme may
help promote the conformational changes that the translocation peptide may undergo
throughout LeuRS’s reaction cycle. Therefore, FN1 and D391A may reduce the rates of
translocation, which impairs the biochemical processes of LeuRS that ultimately leads to
bacterial death.
References
Arnez, J. G., & Moras, D. (1997). Structural and functional considerations of the
aminoacylation reaction. Trends Biochem Sci, 22(6), 211-216.
Betha, A. K., Williams, A. M., & Martinis, S. A. (2007). Isolated CP1 domain of Escherichia
coli leucyl-tRNA synthetase is dependent on flanking hinge motifs for amino acid
editing activity. Biochemistry, 46(21), 6258-6267.
Boniecki, M. T., & Martinis, S. A. (2012). Coordination of tRNA synthetase active sites for
chemical fidelity. J Biol Chem, 287(14), 11285-11289. doi: 10.1074/jbc.C111.325795
Bonnet, J., & Ebel, J. P. (1972). Interpretation of incomplete reactions in tRNA
aminoacylation. Aminoacylation of yeast tRNA Val II with yeast valyl-tRNA
synthetase. Eur J Biochem, 31(2), 335-344.
Cusack, S., Yaremchuk, A., & Tukalo, M. (2000). The 2 A crystal structure of leucyl-tRNA
synthetase and its complex with a leucyl-adenylate analogue. Embo J, 19(10), 23512361.
Drummond, D. A., & Wilke, C. O. (2008). Mistranslation-induced protein misfolding as a
dominant constraint on coding-sequence evolution. Cell, 134(2), 341-352. doi:
10.1016/j.cell.2008.05.042
Fersht, A. R. (1998). Sieves in sequence. Science, 280(5363), 541.
Fersht, A. R., & Dingwall, C. (1979). Evidence for the double-sieve editing mechanism in
protein synthesis. Steric exclusion of isoleucine by valyl-tRNA synthetases.
Biochemistry, 18(12), 2627-2631.
Francklyn, C. S., First, E. A., Perona, J. J., & Hou, Y. M. (2008). Methods for kinetic and
thermodynamic analysis of aminoacyl-tRNA synthetases. Methods, 44(2), 100-118.
doi: 10.1016/j.ymeth.2007.09.007
Fukai, S., Nureki, O., Sekine, S., Shimada, A., Tao, J., Vassylyev, D. G., & Yokoyama, S.
(2000). Structural basis for double-sieve discrimination of L-valine from L-isoleucine
and L-threonine by the complex of tRNA(Val) and valyl-tRNA synthetase. Cell,
103(5), 793-803.
Hartlein, M., & Madern, D. (1987). Molecular cloning and nucleotide sequence of the gene
for Escherichia coli leucyl-tRNA synthetase. Nucleic Acids Res, 15(24), 1019910210.
Bridges 9 (Spring 2015)
12
Hellmann, R. A., & Martinis, S. A. (2009). Defects in transient tRNA translocation bypass
tRNA synthetase quality control mechanisms. J Biol Chem, 284(17), 11478-11484.
doi: 10.1074/jbc.M807395200
Hendrickson, T., & Schimmel, P. (2003). Transfer RNA-Dependent Amino Acid
Discrimination by Aminoacyl-tRNA Synthetases. In J. Lapointe & L. Brakier-Gingras
(Eds.), Translation Mechanisms (pp. 34-64): Eurekah.com and Kluwer
Academic/Plenum Publishers.
Ibba, M., & Soll, D. (2000). Aminoacyl-tRNA synthesis. Annu Rev Biochem, 69, 617-650.
doi: 10.1146/annurev.biochem.69.1.617
Karkhanis, V. A., Boniecki, M. T., Poruri, K., & Martinis, S. A. (2006). A viable amino acid
editing activity in the leucyl-tRNA synthetase CP1-splicing domain is not required in
the yeast mitochondria. J Biol Chem, 281(44), 33217-33225.
Kelly, S. M., Jess, T. J., & Price, N. C. (2005). How to study proteins by circular dichroism.
Biochim Biophys Acta, 1751(2), 119-139. doi: 10.1016/j.bbapap.2005.06.005
Lee, J. W., Beebe, K., Nangle, L. A., Jang, J., Longo-Guess, C. M., Cook, S. A., Ackerman,
S. L. (2006). Editing-defective tRNA synthetase causes protein misfolding and
neurodegeneration. Nature, 443(7107), 50-55.
Lin, L., Hale, S. P., & Schimmel, P. (1996). Aminoacylation error correction. Nature,
384(6604), 33-34.
Lincecum, T. L., Jr., Tukalo, M., Yaremchuk, A., Mursinna, R. S., Williams, A. M., Sproat,
B. S., . . . Cusack, S. (2003). Structural and mechanistic basis of pre- and posttransfer
editing by leucyl-tRNA synthetase. Mol Cell, 11(4), 951-963.
Low, B., Gates, F., Goldstein, T., & Soll, D. (1971). Isolation and partial characterization of
temperature-sensitive Escherichia coli mutants with altered leucyl- and seryl-transfer
ribonucleic acid synthetases. J Bacteriol, 108(2), 742-750.
Mascarenhas, A. P., & Martinis, S. A. (2008). Functional segregation of a predicted "hinge"
site within the beta-strand linkers of Escherichia coli leucyl-tRNA synthetase.
Biochemistry, 47(16), 4808-4816. doi: 10.1021/bi702494q
Mursinna, R. S., Lincecum, T. L., Jr., & Martinis, S. A. (2001). A conserved threonine within
Escherichia coli leucyl-tRNA synthetase prevents hydrolytic editing of leucyltRNALeu. Biochemistry, 40(18), 5376-5381.
Mursinna, R. S., & Martinis, S. A. (2002). Rational design to block amino acid editing of a
tRNA synthetase. J Am Chem Soc, 124(25), 7286-7287.
Nangle, L. A., Zhang, W., Xie, W., Yang, X. L., & Schimmel, P. (2007). Charcot-MarieTooth disease-associated mutant tRNA synthetases linked to altered dimer interface
and neurite distribution defect. Proc Natl Acad Sci U S A, 104(27), 11239-11244.
Bridges 9 (Spring 2015)
13
Normanly, J., Ogden, R. C., Horvath, S. J., & Abelson, J. (1986). Changing the identity of a
transfer RNA. Nature, 321(6067), 213-219.
Palencia, A., Crepin, T., Vu, M. T., Lincecum, T. L., Jr., Martinis, S. A., & Cusack, S.
(2012). Structural dynamics of the aminoacylation and proofreading functional cycle
of bacterial leucyl-tRNA synthetase. Nat Struct Mol Biol, 19(7), 677-684. doi:
10.1038/nsmb.2317
Puglisi, J. D., & Tinoco, I., Jr. (1989). Absorbance melting curves of RNA. Methods
Enzymol, 180, 304-325.
Rao, S. T., & Rossmann, M. G. (1973). Comparison of super-secondary structures in
proteins. J Mol Biol, 76(2), 241-256.
Rock, F. L., Mao, W., Yaremchuk, A., Tukalo, M., Crepin, T., Zhou, H., Alley, M. R. (2007).
An antifungal agent inhibits an aminoacyl-tRNA synthetase by trapping tRNA in the
editing site. Science, 316(5832), 1759-1761. doi: 10.1126/science.1142189
Rould, M. A., Perona, J. J., Soll, D., & Steitz, T. A. (1989). Structure of E. coli glutaminyltRNA synthetase complexed with tRNA(Gln) and ATP at 2.8 A resolution. Science,
246(4934), 1135-1142.
Sampson, J. R., & Uhlenbeck, O. C. (1988). Biochemical and physical characterization of an
unmodified yeast phenylalanine transfer RNA transcribed in vitro. Proc Natl Acad Sci
U S A, 85(4), 1033-1037.
Schmitt, E., Meinnel, T., Blanquet, S., & Mechulam, Y. (1994). Methionyl-tRNA synthetase
needs an intact and mobile 332KMSKS336 motif in catalysis of methionyl adenylate
formation. J Mol Biol, 242(4), 566-576. doi: 10.1006/jmbi.1994.1601
Schreier, A. A., & Schimmel, P. R. (1972). Transfer ribonucleic acid synthetase catalyzed
deacylation of aminoacyl transfer ribonucleic acid in the absence of adenosine
monophosphate and pyrophosphate. Biochemistry, 11(9), 1582-1589.
Silvian, L. F., Wang, J., & Steitz, T. A. (1999). Insights into editing from an ile-tRNA
synthetase structure with tRNAile and mupirocin. Science, 285(5430), 1074-1077.
Starzyk, R. M., Burbaum, J. J., & Schimmel, P. (1989). Insertion of new sequences into the
catalytic domain of an enzyme. Biochemistry, 28(21), 8479-8484.
Swairjo, M. A., & Schimmel, P. R. (2005). Breaking sieve for steric exclusion of a
noncognate amino acid from active site of a tRNA synthetase. Proc Natl Acad Sci U S
A, 102(4), 988-993.
Webster, T., Tsai, H., Kula, M., Mackie, G. A., & Schimmel, P. (1984). Specific sequence
homology and three-dimensional structure of an aminoacyl transfer RNA synthetase.
Science, 226(4680), 1315-1317.
Woese, C. R., Olsen, G. J., Ibba, M., & Soll, D. (2000). Aminoacyl-tRNA synthetases, the
genetic code, and the evolutionary process. Microbiol Mol Biol Rev, 64(1), 202-236.
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Zhao, M. W., Zhu, B., Hao, R., Xu, M. G., Eriani, G., & Wang, E. D. (2005). Leucyl-tRNA
synthetase from the ancestral bacterium Aquifex aeolicus contains relics of synthetase
evolution. Embo J, 24(7), 1430-1439.
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Author
Daniel McDonough graduated from Coastal Carolina University
with a Bachelor’s degree in Chemistry in 2013. He then went on to
pursue a Master’s degree in Business Administration from Coastal
Carolina University. He recently graduated May 2015 with his
MBA. He plans to utilize both degrees as an entrepreneur. Daniel
completed his research project with Dr. Rachel Whitaker during the
academic year 2012-2013.
Adviser/Co-Author
Rachel Whitaker is an Assistant Professor in Biochemistry in the Department
of Chemistry and Physics at Coastal Carolina University, Conway, South
Carolina. She teaches biochemistry and physical biochemistry as well as
general chemistry courses. Her research focuses on enzyme structure and
function with an emphasis on kinetics. Her publications in this area include
one article in the Journal of Biological Chemistry and another in the Journal
of Inorganic Biochemistry.
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